Author Archives: Sarah Gignoux-Wolfsohn


We’re beginning to get data back from a number of experiments, including the Laby culling experiment and Oyster filtration experiment.  At these times, I try to remind myself to remain impartial.  As much as I may want a particular result, and even if negative results are less exciting than positive ones, I just have to wait and let the data speak for themselves.  We have methods for controlling our own bias, having multiple researchers repeat the same tasks, blinding ourselves so we dont see the treatment as were recording results, and ultimately this is the whole point of science.  As Neil deGrasse Tyson said:

Eating your study organism

There is a lot of debate about whether or not it is advisable to eat your organism of study.  I had a professor a few semesters ago who was very adamant about the importance of only studying delicious animals.  He even named this “principle” after himself.

Last weekend, I sampled some oysters from the Island’s only oyster farm, westcott bay shellfish farm.  I enjoyed eating them and also pointing out my newly learned anatomy identification skills to my boyfriend (who did not appreciate this):


Yesterday in class as Carolyn was showing us pictures of diseased oysters, she pointed to a particularly nasty looking one and said “this is why I don’t eat oysters”.  It almost (almost) made me reconsider my decision to adamantly and voraciously eat oysters.

Today in lab, we dissected the baby oysters from the laby experiment to test different tissue types for presence of laby.  We saved samples for histology, pcr, and microscope work.  I couldn’t help but get a little hungry when I shucked the oyster.



What about you? does studying something make you more or less likely to want to eat it?

Primer-dimer, non-specific binding, and other pcr issues

Today, we ran the gel for some of the seastar and Laby PCRs.  In spite of having rerun the PCRs, we got the same results.  The seastar primers seem to bind to non-specific regions and form primer-dimers.


The primer-dimer is the lowest band on this gel.  Primer-dimers are formed by the two primers annealing to each other.  Unfortunately, it was difficult to distinguish from the intended product as we were trying to amplify a 100bp segment, which would be slightly above where the primer-dimer is.  We can also see some non-specific binding in the larger bands.  This can sometimes be fixed by raising the annealing temperature, but in our instance where we weren’t amplifying the intended product, seems to be a sign that we need to reexamine our protocol.  Hopefully we’ll have better luck next time!!

If you’re interested in reading more about troubleshooting PCRs, this guide is very helpful:

Ocean Acidification

Today was spent mostly focused on ocean acidification.  We had a great lecture from carolyn about some of her work looking at transgenerational effects of ocean acidification on oysters.  As the oceans absorb more CO2 and the pH continues to drop, it will be increasingly more important that we understand how acidic conditions affect marine organisms. Another especially interesting aspect of this is the potential of organisms to adapt to these environments.  Her findings suggest that oysters may be able to compensate for some of the deleterious effects of acidic oceans especially if their parents have been exposed to high CO2 conditions.  While these studies are very interesting, controlling CO2 levels and acidity is a very time consuming and difficult process.  Later in the day, we got a tour of the Ocean Acidification lab.


Here, we see the intense filtration process the seawater goes through in order to remove all bacteria and eukaryotes.  Michelle described to us the difficulty in controlling diatom outbreaks and how during an outbreak she would have to change the filters every few hours.


Here we see one of their experimental setups, with pH regulated seawater coming into the mesocosms.  Pretty interesting but also technically complicated stuff!!


PCR Voodoo

When I started my PhD, I spent about three months trying to run a PCR using universal bacterial (16s) primers without a contaminated negative control.  I had run PCRs on more specific genes before, and had never encountered this problem.  I went through everything.  I made new aliquots, I disinfected the entire lab with both ethanol and RNAseAway (which cleaves DNA), I tried switching labs, putting tin foil on the bench, autoclaving my pipets…you name it, I tried it.  Finally I decided the problem was the taq I was using.  It was a green mastermix from a company I wont name and after many hours crying to their technicians on the phone that something must be wrong, I decided to just switch the taq.  Lo and behold, my negative controls became negative.  This experience gave me my nickname (dirty GW) and also initiated me into the world of PCR voodoo.  We have a number of figurines in our lab ranging from pokemon to a picture of tim tebow which are fondly referred to as “PCR gods”.  There is nothing more frustrating than a PCR that doesn’t work, and sometimes your only hope is to pray to whatever gods are available and try random things until something helps.

Today, we did our first PCR of the class on both the seastar and Laby samples.  There are lots of people in our course from different labs.  It’s funny to see the differences in how we do things.  While there are obviously multiple ways to set up a pcr, label your tubes, or run a gel, we all feel shocked by each others methods.  I always label my PCR tubes the same way.  I write a list of samples in my lab notebook, and then label the tubes with corresponding numbers:

photo(6)                           IMG_1185

Other people prefer to write the sample names down the side.  You become so ingrained in your way that another method seems completely absurd.  I always wipe my bench and pipets down with RNAseAway before setting up a PCR.  Lisa uses a 10% bleach solution and has no problem.  Is my sterilization excessive? probably.  but aforementioned traumas make me feel nervous about doing anything less.  When discussing gels, we discovered that the amounts of ethidium bromide we add to the gel varies significantly (I add 0.5ul and Lisa adds 7!!) and the amount of sample and dye we load into the well also varies (2ul-7ul).  How these practices get established can probably be traced down through the scientific lineage and bad or good experiences various people have had with protocols.  For now, We’ll just continue to have nerdy conversations about the absurdity of adding however many microliters of x reagent to a gel.

Culling and Culturing

Today we once again ventured out into the eelgrass beds.  We went to indian cove to look for the plots that had been culled on June 11th.  This time, we were more successful at finding the pvc stakes than we had been at false bay.  There were three transects.  Each transect had four square meter plots.  for each transect, the first plot was a control (not culled) plot the second and third were both culled and the fourth was another control.


We found the center of the plot and placed the quarter meter quadrat around it.  We then sampled every eelgrass shoot in this quarter meter square and brought it back to the lab for analyses.


In the lab, we took the second longest blade of each shoot, measured the length and width, and the size of lesions (if diseased).


With this information, we will be able to see if culling had any impact on prevalence or severity of the disease.

We took some of our classic diseased corals


and placed them on plates to once again culture the laby.


The difficulty of disease signs

Today we surveyed and sampled eelgrass, Zostera marina at False Bay.  Initially, we were attempting to find plots where diseased blades had been culled and blades  had been marked for growth rate in june but the tide was too high.

We did end up running a shallower 50m transect.  We counted blades within a 1/4 m quadrat every 25 m 0m=0blades, 25m=20blades, 50m=0 blades, and collecting samples every 10 meters.  For the collections, we pulled the second longest blade out of 20 bundles.  We then brought these back to the lab to measure and score for disease.  Cody and I had four diseased blades out of our 20:

“classic” eelgrass wasting disease is usually a long lesion surrounded by a dark black border

The disease was difficult to identify, as eelgrass naturally gets lesions.  In order to identify lesions caused by the laby-induced disease, we looked for lesions with a dark black border.  Even this was sometimes difficult to determine, however.

Once we identified diseased and healthy eelgrass, we plated two disease on one plate and two healthy on another.  The lesion or control was cut with sterile scissors, dipped in sterile seawater, blotted, and dipped in 95% ethanol and dried (to remove external bacteria/laby) before being placed on a SSA plate with penicillin and streptomycin.  We will observe these plates in a few days for evidence of laby growth.  Hopefully, we can develop better diagnostic tools for eelgrass wasting disease to reduce some of the difficulty of diagnosis through disease signs alone.


One of the trickiest parts of studying diseases is making sure that you are not cross-contaminating your diseased and healthy samples.  We maintain various levels of sterility based on what we think is necessary and what is possible given our location.  In the lab we may be able to flame sterilize and bleach our instruments in between sampling, but this is not always possible in the field.  Sometimes, our sterility practices almost seem like superstition, rituals we do hoping we are being overly cautious so there is no chance of contamination.  When you have an unknown pathogen, sometimes you only know if your protocols work when you get the results of the experiment.  When it does work, when all our negative controls are negative,  we don’t want to mess with it.  So maybe that second ethanol watch or bleaching isn’t necessary, but it works.


Reyn sterilizing instruments in between sampling Pisaster ochraceus tissue for qPCR.

Drew brought up another important aspect to sterility today when we were surveying seastar wasting syndrome, sterilization of ourselves when we go in between sites.  We bleached our boots after walking among the sick seastars to eliminate the chance of spreading it to other healthier sites.  It’s important to remember that people (and even scientists!) can be disease vectors too, and we should treat ourselves like we treat our sampling equipment.



Today in our lab we looked at the big picture (dissections of molluscs and echinoderms) and the small picture (histology slides of diseased and healthy oysters and abalone).  The big picture allowed us to familiarize ourselves with the various body parts of these animals and also to discover just how hard it is to cut through a pisaster ochracheus.  The small picture introduced us to the difficulties and sometimes joys of differentiating tissue types and identifying parasites.

Here is our dissection of a diseased pisaster ochraceus:


You can see the Anus in the middle surrounded by the stomach, the radial canals/nerves extending down the arms, the yellowish gonads and greenish cecae or digestive glands, and the ampulae (white sacks on each arm) which control the tube feet.

Next, we have a picture of a sectioned and stained abalone infected with Coccidia:


Coccidiae are obligate intracellular parasites, which infect the kidneys of abalone.